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The Journal of Neuroscience, June 15, 2000, 20(12):4423-4434
Prolonged Synaptic Currents and Glutamate Spillover at the
Parallel Fiber to Stellate Cell Synapse
Adam G.
Carter and
Wade G.
Regehr
Department of Neurobiology, Harvard Medical School, Boston,
Massachusetts 02115
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ABSTRACT |
Although neurons often fire in bursts, most of what is known about
glutamate signaling and postsynaptic receptor activation is based on
experiments using single stimuli. Here we examine the activation of
ionotropic glutamate receptors by bursts at the parallel fiber to
stellate cell synapse. We show that brief stimulus trains generate
prolonged AMPA receptor (AMPAR)- and NMDA receptor
(NMDAR)-mediated EPSCs recorded in whole-cell voltage clamp. These
EPSCs contrast with the rapid AMPAR-mediated EPSC evoked by a single
stimulus. The prolonged AMPAR-mediated EPSC is promoted by
high-frequency and high-intensity trains and can persist for hundreds
of milliseconds. This EPSC is also increased by
L-trans-2,4-PDC, an inhibitor of glutamate
transporters, suggesting that these transporters usually limit the
synaptic response to trains. These prolonged EPSCs reflect both
receptor properties and a long-lasting glutamate signal. In addition,
several experiments demonstrate that glutamate spillover can contribute
to receptor activation. First, imaging stimulus-evoked changes in
presynaptic calcium establishes that distinct parallel fiber bands can
be activated. Second, activation of parallel fibers that do not
directly synapse onto a given stellate cell can evoke indirect AMPAR-
and NMDAR-mediated EPSCs in that cell. Third, experiments using the use-dependent NMDAR blocker MK-801 show that these indirect EPSCs reflect glutamate spillover in response to trains. Together, these findings indicate that stimulus trains can generate a sustained and
widespread glutamate signal that can in turn evoke large and prolonged
EPSCs mediated by ionotropic glutamate receptors. These synaptic
properties may have important functional consequences for stellate cell firing.
Key words:
granule cell; parallel fiber; stellate cell; cerebellum; AMPA receptor; NMDA receptor; glutamate; transporter; spillover
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INTRODUCTION |
Neurons often fire in bursts, during
which a variety of processes can enhance or decrease the synaptic
response. Most studies of short-term synaptic plasticity during
repetitive activity have focused on presynaptic processes such as
facilitation and depression (Magleby, 1987 ; Zucker, 1989, 1999).
However, the magnitude and extent of the glutamate signal and the
properties of postsynaptic receptors can also shape the synaptic
response to bursts (Jonas and Spruston, 1994; Clements, 1996; Frerking
and Wilson, 1996; Barbour and Hausser, 1997; Kullmann and Asztely,
1998; Bergles et al., 1999).
Previous studies of postsynaptic receptor activation have often focused
on the synaptic response evoked by a single stimulus. At most
excitatory central synapses, a single stimulus produces a large and
brief glutamate transient in the synaptic cleft (Clements et al., 1992;
Diamond and Jahr, 1997). This glutamate is rapidly removed from the
cleft by diffusion and glutamate transporters. A train of stimuli can
produce much more glutamate release than a single stimulus because
of presynaptic facilitation and delayed release (Atluri and
Regehr, 1996, 1998). The resulting glutamate levels may be sufficiently
large to overwhelm clearance mechanisms, allowing an extended glutamate
signal and even glutamate spillover to nearby sites. Thus, the nature
of the glutamate signal produced by a train may be qualitatively very
different from that generated by a single stimulus.
The importance of an extended glutamate signal and spillover has been
addressed previously in considering the activation of metabotropic
glutamate receptors (mGluRs) (Scanziani et al., 1997; Min et al., 1998;
Vogt and Nicoll, 1999). A stimulus train can activate presynaptic
mGluRs via glutamate spillover to mediate heterosynaptic depression,
whereas a single stimulus is usually ineffective. This consequence of
repetitive activity reflects in part the properties of mGluRs, which
are often located at extrasynaptic sites and have a high affinity for glutamate.
However, the importance of glutamate spillover is less clear for
activation of ionotropic glutamate receptors. At most synapses, a
single stimulus can effectively activate both AMPA receptors (AMPARs) and NMDA receptors (NMDARs) (Hestrin et al., 1990b;
Edmonds et al., 1995). Because the glutamate signal is brief,
deactivation kinetics usually define the synaptic responses mediated by
these receptors (Hestrin, 1992, 1993; Barbour et al., 1994; Edmonds et
al., 1995). Kainate receptors (KARs) can also be activated by a
stimulus train but are poorly activated by a single stimulus (Castillo
et al., 1997; Vignes and Collingridge, 1997). In some cases, a train
may also produce sufficient spillover to activate NMDA receptors at
nearby sites (Kullmann et al., 1996). In contrast, AMPA receptors are
less sensitive to this spillover because they have a lower glutamate
affinity and faster desensitization kinetics (Patneau and Mayer, 1990;
Edmonds et al., 1995). Nevertheless, for synapses at which a large
amount of glutamate is released into a confined space, as at calyceal
and glomerular synapses, a train can produce a sustained elevation of
glutamate and prolonged AMPA receptor activation (Rossi et al., 1995;
Otis and Trussell, 1996; Otis et al., 1996; Silver et al., 1996; Kinney
et al., 1997; Slater et al., 1997; Overstreet et al., 1999). However,
most synapses lack this specialized anatomy, and glutamate spillover is
generally not thought to activate AMPA receptors.
Here we study the synaptic response to brief stimulus trains at the
parallel fiber to stellate cell synapse in the transverse cerebellar
slice of young rats. This preparation allows us to stimulate distinct
parallel fiber inputs to a given stellate cell and to record the evoked
synaptic response using whole-cell voltage clamp. We find that
high-frequency and high-intensity trains yield a large and prolonged
EPSC in stellate cells. We show that this EPSC is mediated by both AMPA
and NMDA receptors and reflects an extended glutamate signal which
itself partly reflects glutamate spillover at this synapse.
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MATERIALS AND METHODS |
Transverse cerebellar slices (300-µm-thick) were cut from 17- to 21-d-old Sprague Dawley rats, as described previously (Llano et al.,
1991; Atluri and Regehr, 1996). Experiments were conducted at either
24 ± 1 or 34 ± 1°C. The control external solution
consisted of (in mM): 125 NaCl, 2.5 KCl, 1.5 CaCl2, 1 MgCl2, 26 NaHCO3, 1.25 NaH2PO4, 25 glucose, and
0.02 bicuculline, bubbled with 95% O2/5%
CO2. Flow rates at 24 and 34°C were 2-3 and
4-6 ml/min, respectively.
Parallel fibers were stimulated using an extracellular glass
microelectrode placed in the molecular layer  100 µm from the recording electrode. The tip diameter of this stimulus electrode was
typically 10 µm, and stimulus intensities ranged from 5 to 40 µA
with stimulus durations of 0.1-0.5 msec. In some experiments, a second
stimulating electrode was placed in the molecular layer tens of
micrometers from the first electrode, and two tracts of parallel
fibers were independently stimulated.
All recordings were made from stellate cells in the outer two-thirds of
the molecular layer of the cerebellar cortex. EPSCs were recorded using
whole-cell voltage clamp with an Axopatch 200B amplifier (Axon
Instruments, Foster City, CA). These recordings were obtained
using 1.8-2.5 M glass pipettes containing an internal solution of
(in mM): 35 CsF, 100 CsCl, 10 EGTA, and 10 HEPES, pH
7.3-7.4. Series resistance (typically 5-10 M) and leak current (typically  5 to 20 pA at 40 mV, and 10 to 30 pA at 70 mV) were monitored continuously, and experiments were rejected if either
value worsened. Pipettes were wrapped in parafilm to reduce pipette
capacitance, as was also the case for the other recording configurations described below. No series resistance compensation was
used. Unless otherwise noted, cells were held at 0 mV between parallel
fiber stimulations.
Synaptic charge (EPSCcharge) was the integrated
synaptic current. For single stimuli, the integration period began
immediately after the prespike and extended for 10-50 msec. For
trains, the integration period began immediately after the last
stimulus. Because of the variability of the slow component, the
integration time was chosen to match the time course of the current and
ranged from 0.5 to 2 sec. Errors are expressed as SEM.
Stellate cell firing was monitored using both cell-attached patch and
whole-cell current-clamp recordings. Cell-attached patch recordings
were obtained using 3-4.5 M glass pipettes containing a pipette
solution of (in mM): 152.5 NaCl, 2.5 KCl, 1.5 CaCl2, 1 MgCl2, 10 HEPES,
and 10 glucose, pH 7.3-7.4. Whole-cell current-clamp recordings were
obtained using 3-4.5 M glass pipettes containing an internal
solution of (in mM): 130 KMeSO3, 10 NaCl, 2 MgCl2, 0.5 EGTA, 10 HEPES, 4 MgATP, 14 creatine phosphate, and 0.3 GTP, pH 7.3-7.4. Current-clamp recordings
were made in fast I-clamp mode on the Axopatch 200B amplifier.
All chemicals were from Sigma (St. Louis, MO) with the exception of
MCPG [(RS)- -methyl-3-carboxymethylphenylglycine], CPPG [(RS)--cyclopropyl-4-phosphonophenylglycine], NBQX
(2,3-dioxo-6-nitro-1,2,3,4-tetrahydrobenzo[f]quinoxaline-7-sulfonamide), D-AP-5
(D()-2-amino-5-phosphonopentanoic acid), PDC
(L-trans-2,4-PDC), CTZ
[6-chloro-3,4,-dihydro-3-(2-norbornen-5-yl)-2H-1,2,4-benzothiadiazine- 7-sulfonamide-1,1-dioxide;
cyclothiazide], MK-801
[(5R,10S)-(+)-5- methyl-10,11-dihydro-5H-dibezo[a,d]cyclohepten-5,10-imine]
(Tocris Cookson, Ballwin, MO), and GYKI 53655 (gift from Eli Lilly & Co., Indianapolis, IN).
Outputs of the Axopatch 200B were filtered at 5 kHz and digitized with
a 16 bit digital-to-analog converter (Instrutech, Great Neck, NY),
Pulse Control software (Herrington and Bookman, 1995), and an Apple
Computers (Cupertino, CA) Macintosh Quadra 650 computer. Analysis was
done on- and off-line with Igor Pro software (WaveMetrics, Lake Oswego, OR).
For the experiment shown in Figure 7, parallel fibers were labeled
using local application of the membrane-permeant dye Oregon Green 488 BAPTA-1 AM (Molecular Probes, Eugene, OR) in solution, as described
previously (Regehr and Tank, 1991; Regehr and Atluri, 1995). The
loading time was 20 min, and the imaging was performed ~2 hr later.
Two bands of parallel fibers were stimulated using pipettes with tip
diameters of 10 µm, with a stimulus intensity of 5 µA and stimulus
duration of 0.5 msec. Fluorescence was detected using an Olympus
Optical (Tokyo, Japan) confocal microscope equipped with an argon
laser. Scan time for each image was ~1 sec.
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RESULTS |
Synaptic responses to brief stimulus trains were studied at the
granule cell to stellate cell synapse in transverse cerebellar brain
slices of young rats. In these slices, granule cell parallel fibers run
across the surface of the molecular layer where they contact several
types of neurons (Palay and Chan-Palay, 1974). One such target is the
stellate cell, a small inhibitory interneuron located in the outer
two-thirds of the molecular layer. We stimulated parallel fibers with
glass microelectrodes placed in the molecular layer and recorded EPSCs
in stellate cells using whole-cell voltage clamp. All experiments were
performed in the presence of bicuculline to eliminate synaptic
responses mediated by GABAA receptors.
Stimulus trains and the prolonged synaptic response
As shown previously, a single stimulus evoked a fast EPSC in a
stellate cell held at 60 mV (Fig.
1A) (Barbour et al.,
1994; Atluri and Regehr, 1998). The fast decay of this EPSC reflects a
rapid glutamate signal and AMPAR deactivation (Barbour et al., 1994).
In contrast, a brief stimulus train of five pulses at 100 Hz generated
a more complex synaptic response (Fig. 1B). The peak EPSCs of this response increased during the train, reflecting an
increase in the probability of release caused by facilitation (Atluri
and Regehr, 1998). In addition, the EPSC had a prolonged component that
is more difficult to explain. The remainder of the paper details
experiments aimed to clarify the nature of this prolonged EPSC.
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Figure 1.
Stellate cell EPSCs evoked by parallel fiber
stimulation. Whole-cell voltage-clamp recordings from stellate cells in
response to one (A) or five
(B) pulses delivered to the parallel fibers at
100 Hz. Recordings were made at 60 mV in the absence of antagonists.
Stimulus artifacts are blanked for clarity, and traces are single trial
examples.
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Pharmacology of the prolonged synaptic response
Previous studies at this synapse have found that synaptic currents
evoked by single stimuli are mediated primarily by AMPARs (Barbour et
al., 1994; Glitsch and Marty, 1999). However, stellate cells are
thought to possess a variety of other glutamate receptors, including
mGluRs (Baude et al., 1993), NMDARs (Monyer et al., 1994; Cull-Candy et
al., 1998), and KARs (Bahn et al., 1994). A stimulus train may produce
a glutamate signal sufficient to activate these receptors, thereby
generating the prolonged EPSC. We thus used pharmacology to determine
the glutamate receptors responsible for mediating the prolonged EPSC
evoked by a stimulus train.
We first used the mGluR antagonists MCPG (group I/II) and CPPG
(group II/III) to determine a role for these receptors in mediating the
prolonged EPSC. We quantified the effects of these antagonists using
the peak synaptic current (EPSCpeak) and the
synaptic charge (EPSCcharge), which is the
integrated synaptic current after the last stimulus in a train (see
Materials and Methods). We found that application of either MCPG
(0.5-1 mM) or CPPG (30 µM) had no effect on
EPSCcharge or EPSCpeak
recorded while holding cells at 40 mV [for MCPG,
EPSCcharge was 99 ± 29% and
EPSCpeak was 95 ± 7% (n = 3) of control; for CPPG, EPSCcharge was 104 ± 26% and EPSCpeak was 92 ± 5%
(n = 3) of control]. Thus, activation of mGluRs does
not contribute to the synaptic response evoked by a brief stimulus
train at this synapse under our recording conditions.
We next used blockers of different ionotropic glutamate receptors to
determine their relative contributions to the synaptic response evoked
by a stimulus train. Coapplication of NBQX (10 µM) and
D-AP-5 (200 µM), blockers of non-NMDARs and
NMDARs, respectively, eliminated this synaptic response at all holding
potentials (data not shown, n = 5). Application of NBQX
alone eliminated the peak EPSCs and revealed an outwardly rectified
EPSC typical of NMDARs (Fig.
2Ai, B).
This EPSC gradually rose during the stimulus train but was usually
small or absent after a single stimulus. However, the small size of
this EPSC at 60 mV suggests that it is not responsible for the
prolonged EPSC shown in Figure 1. Application of
D-AP-5 alone eliminated this outwardly rectified
EPSC and revealed a prolonged EPSC at all holding potentials (Fig.
2Aii, B). Thus, we found that both NMDARs
and non-NMDARs can contribute to the synaptic response to a
stimulus train but that the prolonged EPSC shown in Figure 1 is most
likely mediated by non-NMDARs.
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Figure 2.
Pharmacology of the EPSCs. A, EPSCs
evoked by five pulses at 100 Hz measured at holding potentials of 80,
60, 40, 20, 0, +20, and +40 mV. Recordings were made in the
presence of either 10 µM NBQX (i)
or 200 µM D-AP-5 (ii). B,
Normalized peak EPSCs versus holding potential in the presence of NBQX
(closed circles; n = 3) or
D-AP-5 (open circles; n = 3). Peak EPSCs evoked at different holding potentials were normalized
to those currents evoked at +40 mV. C, EPSCs evoked by
five pulses at 100 Hz measured at 40 mV while in the presence of
either 50 µM D-AP-5 or 30 µM
GYKI 53655 plus 50 µM D-AP-5. Recordings in
A-C are from different cells, and traces are averages
of two to five trials.
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The prolonged non-NMDAR-mediated EPSC could be attributable to
activation of AMPARs or KARs. Synaptic currents mediated by KARs have a
slow time course and can become prominent during stimulus trains,
making KARs a good candidate for contributing to the synaptic response
(Castillo et al., 1997; Vignes and Collingridge, 1997). We tested for
this possibility using GYKI 53655, a selective blocker of AMPARs that
does not block KARs. Application of GYKI 53655 (30 µM)
eliminated the prolonged non-NMDAR-mediated EPSC recorded while holding
cells at 40 mV (EPSCcharge was 6 ± 2%
and EPSCpeak was 5 ± 2% of control;
n = 3) (Fig. 2C). This finding indicates that this EPSC is mediated solely by activation of AMPARs. This is
intriguing, because AMPAR-mediated EPSCs evoked by a single stimulus are very fast at this synapse and these AMPARs rapidly desensitize in the sustained presence of glutamate (Barbour et al.,
1994).
Stimulus conditions that generate the prolonged
AMPAR-mediated EPSC
We next sought to clarify the stimulus conditions that generate
the prolonged AMPAR-mediated EPSC. As indicated in Figure 1, when the
number of pulses in the train was increased from one to five, the EPSC
decayed much more slowly (Fig.
3A). Furthermore, when the
frequency of a five pulse train was increased from 10 to 100 Hz, the
EPSC was again prolonged (Fig. 3B). This is shown by a
scaled comparison of the two EPSCs after the last stimulus in each of
the trains (Fig. 3B, bottom). Moreover, when the
intensity of a five pulse train was increased 10-fold, from 3 to 30 µA, the EPSC again decayed more slowly (Fig. 3C). Thus,
the prolonged AMPAR-mediated EPSC is accentuated by trains of
high-frequency and high-intensity stimuli, and we therefore used such
stimuli throughout the remainder of this study.
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Figure 3.
Stimulus conditions that elicit the prolonged
AMPAR-mediated EPSC. A, EPSCs evoked by one, two, and
five pulses at 100 Hz. B, EPSCs evoked by five pulses at
10 (top) or 100 (middle) Hz, and a scaled
comparison of the time course of decay after the fifth stimulus
(bottom). C, EPSCs evoked by five pulses
at 100 Hz and either 3 (top) or 30 (middle) µA, and a comparison of the responses scaled
to the last peak (bottom). Recordings were made at 40
mV, and 50 µM D-AP-5 was present for all
experiments. Recordings in A-C are from different
cells, and traces are averages of 10-40 trials.
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The prolonged AMPAR-mediated EPSC is not attributable to poor
voltage clamp
Inadequate voltage clamp could contribute to the slow,
non-exponential decay of the prolonged EPSC (Spruston et al., 1993). We
performed two types of experiments to show that this was not the case.
First, we conducted a voltage jump experiment (Hestrin et al., 1990a;
Pearce, 1993; Barbour et al., 1994) in which the cell was stimulated
while held at 0 mV, and the holding potential was stepped to 40 mV at
different times after the stimulus train (Fig.
4Ai). If the prolonged
AMPAR-mediated EPSC reflects poor voltage clamp, there should be
a clear difference between these responses and those evoked while the
cell was held at 40 mV throughout. However, the responses closely
align after returning to 40 mV, indicating that voltage clamp was
adequate (Fig. 4Aii). Similar results were found in
all five cells tested. Next, we applied a low concentration of NBQX
(250 nM) to decrease the amplitude of the EPSC,
which should reduce any effects of poor voltage clamp (Fig.
4B). A scaled comparison of the EPSCs evoked in the
presence and absence of low NBQX shows that a prolonged AMPAR-mediated EPSC is found in both conditions, again indicating that voltage clamp
was adequate (Fig. 4B, bottom). Similar
results were found in all four cells tested.
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Figure 4.
The prolonged AMPAR-mediated EPSC is not a
consequence of poor voltage clamp. A, EPSCs evoked by
five pulses at 100 Hz measured either at 40 mV or after stepping from
0 to 40 mV during the slow component of the EPSC, as shown in the
schematic (i). After the step to 40 mV, the
EPSCs are closely aligned (ii). The capacitative current
has been subtracted. B, EPSCs evoked by five pulses at
100 Hz measured at 40 mV while in the presence of either
D-AP-5 (top; bottom,
thin line) or 250 nM NBQX plus
D-AP-5 (middle; bottom,
thick line), and a comparison of the responses scaled to
the last peak (bottom). D-AP-5 (50 µM) was present for all experiments. Recordings in
A and B are from different cells, and
traces are averages of 6-40 trials.
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Contributions of glutamate transporters and desensitization to the
prolonged AMPAR-mediated EPSC
We next tested the role of glutamate transporters in shaping the
prolonged AMPAR-mediated EPSC evoked by trains. A variety of glutamate
transporters are present in the molecular layer of the cerebellum
(Rothstein et al., 1994; Chaudhry et al., 1995; Lehre et al., 1995). We
blocked these transporters with PDC and assessed the effects on
AMPAR-mediated EPSCs. PDC (200 µM) decreased the EPSC
evoked by a single stimulus (EPSCcharge was
58 ± 13% and EPSCpeak was 55 ± 8%
of control; n = 4) (Fig.
5A). PDC also increased the
leak current, despite the presence of 1 mM
external Mg2+ and
D-AP-5 to block NMDARs (Fig. 5B,
bottom inset). These findings suggest that blocking
glutamate transporters with PDC can increase ambient extracellular
glutamate to concentrations that can activate and subsequently
desensitize a fraction of AMPARs. However, the negligible effect of PDC
on the half-decay time of the EPSC evoked by a single stimulus
(t1/2 was 95 ± 10% of control;
n = 4) indicates that glutamate transporters play a
limited role in shaping this response, in agreement with previous
studies at other synapses (Isaacson and Nicoll, 1993; Sarantis et al.,
1993). In contrast, PDC greatly extended the time course of the
prolonged AMPAR-mediated EPSC evoked by a stimulus train
(EPSCcharge was 560 ± 90%,
EPSCpeak was 96 ± 8%, and
t1/2 was 1020 ± 120% of
control; n = 4) (Fig. 5B). These results
suggest that the prolonged AMPAR-mediated EPSC evoked by a stimulus
train reflects an extended glutamate signal that is controlled by
glutamate uptake.
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Figure 5.
Inhibition of glutamate transporters enhances the
prolonged AMPAR-mediated EPSC. EPSCs evoked by one
(A) and five (B) pulses at
100 Hz while in the presence of either D-AP-5 (thin
line) or 200 µM PDC plus D-AP-5
(thick line). Insets show the effect of
PDC on the scaled synaptic charge transfer or leak current. Recordings
were made at 40 mV, and 50 µM D-AP-5 was
present for all experiments. Recordings in A and
B are from different cells, and traces are averages of
10 trials.
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We also tested the role of AMPAR desensitization in shaping the
response to a stimulus train. We found that 40 µM CTZ,
which reduces AMPAR desensitization, greatly increased the magnitude and time course of the prolonged AMPAR-mediated EPSC evoked by a
stimulus train (EPSCcharge was 970 ± 10%
and EPSCpeak was 370 ± 70% of control;
n = 3) (Fig.
6A). This dramatic
effect of CTZ is consistent with the sustained presence of glutamate
after a train and with AMPAR desensitization limiting the synaptic
response. However, CTZ can also affect the presynaptic probability of
release, AMPAR deactivation, and AMPAR affinity for glutamate (Patneau et al., 1993; Diamond and Jahr, 1995; Dzubay and Jahr, 1999). We found
that CTZ had no significant effect on the NMDAR-mediated EPSC evoked by
a stimulus train (EPSCcharge was 93 ± 5%
and EPSCpeak was also 93 ± 5% of control;
n = 3) (Fig. 6B). Because CTZ does not directly affect NMDAR kinetics or glutamate affinity (Trussell et
al., 1993; Mennerick and Zorumski, 1995), we conclude that CTZ does not
have significant presynaptic effects under our experimental conditions.
We also found that CTZ had a much smaller effect on the AMPAR-mediated
EPSC evoked by a single stimulus (EPSCcharge was
380 ± 50% and EPSCpeak was 220 ± 30% of control; n = 5) (Fig. 6C) compared
with the AMPAR-mediated EPSC evoked by a train. This suggests that,
although CTZ affects AMPAR deactivation or glutamate affinity, the
effect of CTZ on the synaptic response to a stimulus train primarily
reflects the reduction of AMPAR desensitization caused by the prolonged
elevation of glutamate.
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Figure 6.
Reduction of AMPAR desensitization enhances the
prolonged AMPAR-mediated EPSC. A, AMPAR EPSCs evoked by
five pulses at 100 Hz measured at 40 mV while in the presence of
either 50 µM D-AP-5 or 40 µM
CTZ plus 50 µM D-AP-5. B,
NMDAR EPSCs evoked by five pulses at 100 Hz while in the presence of
either 10 µM NBQX or 40 µM CTZ plus 10 µM NBQX. C, AMPAR EPSCs evoked by one
pulse measured at 40 mV while in the presence of either 50 µM D-AP-5 or 40 µM CTZ plus 50 µM D-AP-5. Insets show the
effect of CTZ on the scaled synaptic charge transfer. Recordings in
A-C are from different cells, and traces are averages
of 4-10 trials.
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Imaging activated bands of parallel fiber
The above findings suggest that trains can produce a sustained
elevation of glutamate and raise the possibility that glutamate spillover occurs at this synapse. We next tested for glutamate spillover by taking advantage of the anatomy of the parallel fibers. The goals of these experiments were (1) to determine whether
stimulating a pathway that does not directly contact a stellate cell
can evoke a synaptic response in that cell, and (2) to show that this
response reflects the activation of receptors located at nearby
synapses via glutamate spillover. For such experiments, it is necessary to stimulate well defined bands of parallel fibers.
We imaged stimulus-evoked changes in presynaptic calcium to demonstrate
our ability to stimulate distinct bands of parallel fibers. Parallel
fibers were labeled with the calcium-sensitive indicator Oregon Green
488 BAPTA-1 AM (Fig. 7A) (see
Materials and Methods). The resulting fluorescence revealed that
parallel fibers were labeled, but other cellular structures were not
(Fig. 7B). Two stimulus electrodes (S1 and S2) were then
placed ~80 µm apart in the molecular layer, and two distinct bands
of parallel fibers were stimulated (100 pulses at 100 Hz). Stimulation
of S1 produced an increase in fluorescence in a band of fibers that is
apparent in the raw fluorescence (Fig. 7C) and in a trace
for which the baseline fluorescence has been subtracted (Fig.
7E). Stimulation with S2 produced similar results in a
separate band of fibers (Fig.
7D,F). These fluorescence
changes arise from increases in intracellular
Ca2+ levels, and they provide a good
measure of the spatial extent of fiber activation. For stimulation with
either S1 or S2, the activated bands of fibers were ~40 µm in
diameter, and these bands did not overlap. Similar stimulation of bands
of fibers was observed in all six slices tested.
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Figure 7.
Calcium imaging of parallel fiber bands.
A, Schematic illustrating the load site, labeled fibers,
location of stimulus electrodes S1 and S2, and field of view in
B-F. B, Background fluorescence of
fibers labeled with Oregon Green 488 BAPTA-1 AM. C,
D, Fluorescence evoked by 100 pulses at 100 Hz for
electrode S1 (C) or electrode S2
(D). E, F, Evoked
changes in fluorescence for S1 (E) and S2
(F) in which B has been subtracted
away from C and D, respectively.
Traces are single trial examples.
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Indirect EPSCs
If glutamate spillover occurs at the parallel fiber to stellate
cell synapse, then activation of presynaptic terminals that do not
directly contact a stellate cell might give rise to a detectable synaptic response. We tested for this possibility by examining EPSCs
produced by activation of two parallel fiber pathways, with the goal to
identify a direct pathway that makes synaptic contacts onto a
particular stellate cell and an indirect pathway that does not (Fig.
8A). The anatomy of the
parallel fibers and our imaging of activated fiber tracts (Fig. 7)
indicate that it should be possible to stimulate these two pathways. In
the presence of D-AP-5, stimulation of one
pathway (five pulses at 100 Hz) yielded fast AMPAR-mediated EPSCs,
followed by the prolonged AMPAR-mediated EPSC (Fig.
8Bi). Based on the presence of the prominent fast
EPSCs, this was identified as a direct pathway. In contrast,
stimulation of another pathway (20 pulses at 100 Hz) failed to elicit a
significant fast response (compare the first 50 msec after the onset of
stimulation in Fig. 8Bi, Bii) but did
produce a small EPSC that gradually developed and slowly decayed (Fig.
8Bii, Biii). Based on the lack of fast
EPSCs, this second pathway was identified as an indirect pathway. This
indirect response was blocked by 10 µM NBQX
(EPSCcharge was 0.6 ± 0.6% of control;
n = 3) (Fig. 8Biii) and 30 µM GYKI 53655 (EPSCcharge
was 3.1 ± 1.5% of control; n = 5) (data not shown), suggesting that it was mediated by AMPARs. This indirect response was greatly enhanced by 200 µM PDC
(EPSCcharge was 2100 ± 700% of control;
n = 4) (Fig. 8C), indicating that glutamate transporters usually limit this response. This indirect AMPAR-mediated response was also increased by 40 µM CTZ
(EPSCcharge was 1050 ± 80% of control;
n = 3) (data not shown), suggesting that AMPAR desensitization usually limits this response.
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Figure 8.
AMPAR- and NMDAR-mediated EPSCs evoked by indirect
pathway stimulation. A, Schematic illustrating direct
(S1) and indirect (S2) pathways. B, AMPAR EPSCs evoked
by stimulating a direct pathway with five pulses at 100 Hz
(i) or an indirect pathway with 20 pulses at 100 Hz (ii, iii). Recordings were made at
40 mV, and 50 µM D-AP-5 was present.
Addition of 10 µM NBQX blocked the indirect response
(iii). C, AMPAR EPSCs evoked by five
pulses at 100 Hz measured at 40 mV while in the presence of 50 µM D-AP-5 or 200 µM PDC plus 50 µM D-AP-5. D, NMDAR EPSCs
evoked by stimulating a direct pathway with five pulses at 100 Hz
(i) or an indirect pathway with 20 pulses at 100 Hz (ii, iii). Recordings were made at +40
mV, and 10 µM NBQX was present. Addition of 4 µM MK-801 blocked the indirect response
(iii). The duration of stimulation is indicated by the
horizontal bars. Scale bar in Bi applies
to Bi and Bii; scale bar in
Di applies to Di and Dii.
Recordings in B-D are from different cells, and traces
are averages of four to five trials.
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We next examined indirect synaptic responses mediated by NMDARs, whose
higher glutamate affinity may allow them to better detect glutamate
spillover (Patneau and Mayer, 1990; Edmonds et al., 1995). Direct and
indirect pathways were first identified based on AMPAR-mediated EPSCs.
We then examined the NMDAR-mediated EPSCs in isolation by including
NBQX in the external solution (Fig. 8D). Stimulation
of the direct pathway (five pulses at 100 Hz) to a cell held at +40 mV
yielded an EPSC that began shortly after the first stimulus, peaked
tens of milliseconds after the fifth stimulus, and decayed over several
hundred milliseconds (Fig. 8Di). Stimulation of the
indirect pathway with 20 pulses at 100 Hz yielded a much slower EPSC
that typically began well into the stimulus train, peaked several
hundred milliseconds after the last stimulus, and decayed over several
seconds (Fig. 8Dii, Diii). This indirect
response was blocked by 4 µM MK-801, an
open-channel blocker of NMDARs (EPSCcharge was
1.7 ± 2.5% of control; n = 7) (Fig.
8Diii). The simplest explanation for these indirect
NMDAR- and AMPAR-mediated EPSCs is that glutamate released at synapses during a stimulus train can spillover and activate ionotropic glutamate
receptors on nearby stellate cells.
Indirect NMDAR-mediated EPSC and glutamate spillover
We next used MK-801 to further examine the contribution of
glutamate spillover to synaptic responses evoked by stimulus trains. MK-801 has been a valuable tool in the study of synaptic transmission by virtue of its ability to block only open NMDARs (Huettner and Bean,
1988; Jahr, 1992; Hessler et al., 1993; Rosenmund et al., 1993). We
used MK-801 to block NMDARs opened during the synaptic response to
indirect pathway stimulation. If this also reduces the response to
direct pathway stimulation at the same cell, then the indirect
NMDAR-mediated EPSC must reflect glutamate spillover.
In these experiments, two stimulus electrodes were positioned as in
Figure 8A, such that one evoked an indirect response
(Fig. 9Ai) and the other a
direct response (Fig. 9Aii, larger trace). After
obtaining stable recordings for both pathways, we washed 4 µM MK-801 into the bath. Between successive
stimulation, the cell was held at 70 mV to avoid NMDAR activation by
extracellular glutamate and subsequent block by MK-801. The indirect
pathway was then stimulated once every 30 sec for 5 min (Fig.
9Aiii). During this time, the indirect response
progressively decreased (data not shown). We then tested the response
to stimulation of the direct pathway. For the cell in Figure
9A, the first direct response in MK-801 was 6% of the
average direct response in control conditions (Fig. 9Aii,
smaller trace, Aiii). For seven such experiments, the average amplitude of the first direct response in MK-801 was 33 ± 9% of control. This large decrease suggested that the
indirect response reflects glutamate spillover.
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Figure 9.
The indirect NMDAR-mediated EPSC is a consequence
of glutamate spillover. Ai, Indirect NMDAR EPSC evoked
by 20 pulses at 100 Hz. Aii, Direct NMDAR EPSCs evoked
by five pulses at 100 Hz in control conditions (larger
trace) and in MK-801 (smaller trace, *).
Aiii, Peak direct NMDAR EPSCs as a function of time.
After obtaining a stable recording (circles), 4 µM MK-801 was added to the bath (horizontal dashed
line). The indirect pathway was stimulated at the indicated
times (vertical lines), and then the direct response was
tested. The first time the direct response was tested in MK-801 (*),
the EPSC was greatly reduced in size. In B, the
experiment was repeated, except now the indirect pathway was not
stimulated in the presence of MK-801. Bi, Direct NMDAR
EPSCs evoked by five pulses at 100 Hz in control conditions
(larger trace) and in MK-801 (smaller
trace,  ). Bii, Peak direct NMDAR EPSCs as a
function of time. The first time the direct response was tested in
MK-801 (), the EPSC was only slightly reduced in size. In all of the
experiments, 10 µM NBQX was present, and the holding
potential was stepped from 70 to +40 mV to assess the NMDAR EPSC.
Recordings in A and B are from different
cells.
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We also performed control experiments to verify that the decrease in
the first direct response was attributable to glutamate spillover.
These experiments were identical to those described above, except that
the indirect pathway was not stimulated in the presence of MK-801. We
found that the first direct response in MK-801 was slightly smaller
than the average direct response in control conditions (Fig.
9Bi, Bii). For the cell in Figure 9B,
the first direct response in MK-801 was 85% of control. For seven such
experiments, the average first direct response in MK-801 was 67 ± 7% of control. Much of this decrease likely reflects the development
of MK-801 open-channel block during the first direct response. In
addition, MK-801 could block NMDARs opened by ambient extracellular
glutamate during steps from 70 to +40 mV. Finally, gradual rundown of
the direct response during the 10 min for which it could not be tested
may also contribute to this decrease.
Thus, the decrease in the first direct response in MK-801 after
indirect pathway stimulation is primarily a reflection of glutamate
spillover, because this decrease was much smaller when the indirect
pathway was not stimulated (33 ± 9% with stimulation vs 67 ± 7% without). These experiments thus establish that, at 24°C,
glutamate can diffuse from nearby synapses to activate NMDARs on a
stellate cell.
Prolonged AMPAR-mediated EPSC and glutamate spillover
at 34°C
Previous studies have indicated that glutamate spillover can be
much less prominent at more physiological temperatures (Asztely et al.,
1997). We therefore tested whether the prolonged AMPAR-mediated EPSC
and glutamate spillover that we have described at 24°C are also
present at 34°C.
We found that high-frequency, high-intensity direct pathway stimulation
could generate a prolonged AMPAR-mediated EPSC at 34°C (Fig.
10A). A low-intensity
stimulus train of five pulses at 100 Hz evoked rapidly decaying fast
EPSCs (Fig. 10A, top). However, at a
higher intensity, the same stimulus train produced fast EPSCs and a
prolonged EPSC (Fig. 10A, middle). This is
shown by a scaled comparison of the two synaptic responses (Fig.
10A, bottom). We also found that indirect
pathway stimulation could evoke indirect AMPAR-mediated EPSCs at 34°C
(Fig. 10B). Direct (Fig. 10Bi) and indirect (Fig. 10Bii) pathways were identified based
on the presence of fast EPSCs. The indirect response was blocked by 10 µM NBQX (Fig. 10Biii),
suggesting that it was mediated by AMPARs. These findings suggest that
stimulus trains can generate sustained glutamate signals and glutamate
spillover at 34°C.
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Figure 10.
Slow direct and indirect AMPAR-mediated EPSCs are
present at 34°C. A, Direct AMPAR EPSCs evoked by five
pulses at 100 Hz at low (top) or high
(middle) stimulus intensity and a comparison of the
responses scaled to the last peak (bottom).
B, Direct AMPAR EPSCs evoked by five pulses at 100 Hz
(S1) (i) and indirect AMPAR EPSCs evoked by 20 pulses at 100 Hz (S2) (ii). Addition of 10 µM NBQX blocked the indirect response
(iii). Recordings were made at 40 mV, and 50 µM D-AP-5 was present for all experiments.
Scale bar in Bi applies to Bi and
Bii. Recordings in A and B
are from different cells, and traces are averages of 5-20
trials.
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We also used MK-801 to assess the ability of glutamate spillover to
activate NMDARs at 34°C (Fig. 11). We
found that the decrement of the first direct response in MK-801
depended on whether the indirect pathway was stimulated. In the
examples shown, the first direct response in MK-801 was 18% of control
when the indirect pathway was stimulated (Fig. 11A)
and 74% of control when the indirect pathway was not stimulated (Fig.
11B). On average, the first direct response was
29 ± 10% (n = 4) of control with indirect
pathway stimulation and 61 ± 5% (n = 5) of
control without.
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Figure 11.
The indirect NMDAR-mediated EPSC at 34°C
reflects glutamate spillover. These experiments were the same as those
performed at 24°C (Fig. 9), except that now stimuli were separated by
15 sec, the MK-801 wash in time was 3 min, indirect pathway stimulation
took place over a 3 min period, and the temperature was 34°C.
Recordings in A and B are from different
cells.
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Together, these experiments establish that glutamate levels can be
elevated for hundreds of milliseconds after a stimulus train at 34°C.
Moreover, glutamate can spill over from synapses and activate both
AMPARs and NMDARs at nearby synapses even at this temperature.
Effects of stimulus trains on stellate cell firing
We next examined the effect of direct pathway stimulation on
stellate cell firing at 24°C. We first recorded from stellate cells
using the cell-attached patch configuration, which does not alter the
internal environment of the cells (Fig.
12A). For our
experimental conditions (GABAA receptors are
blocked), these cells are spontaneously active at frequencies of 2-10
Hz. For low-intensity stimulation, a single stimulus did not
consistently evoke a response (Fig. 12Ai,
left), as indicated by the representative trace and the
raster plots for five trials. In contrast, a train of five stimuli at
the same intensity increased stellate cell firing for ~100 msec (Fig.
12Ai, right). At a higher intensity, a
single stimulus could reliably trigger an action potential (Fig. 12Aii, left). Furthermore, a stimulus
train generated a more complex response (Fig. 12Aii,
right) in which firing first briefly increased, then ceased
for hundreds of milliseconds, and subsequently returned to baseline
only after a rebound to relatively high firing rates. Similar effects
on firing were observed in all seven cells tested. The long duration of
the changes in firing after brief trains (Fig. 12Ai,
Aii, right) suggests a role for the prolonged
EPSCs recorded in whole-cell voltage clamp. The response to
high-intensity stimulus trains is particularly interesting in that
activation of an excitatory input paradoxically inhibits stellate cell
firing.
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Figure 12.
Response of stellate cells to parallel fiber
stimulation. Cell-attached patch recordings (A)
and whole-cell current-clamp recordings (B) from
stellate cells in response to one (left) or five (right) pulses at 100 Hz using low- (i) or high- (ii)
intensity stimulation. The high and low stimulus intensities differed
by a factor of 2.5. Beneath each example in A and
B is a raster plot showing spike timing for five
consecutive trials. Below each raster plot is a marker
indicating the time of stimulation. In B and
C, the horizontal markers correspond to
65 and 0 mV. In C, the responses to five pulses at 100 Hz are replotted on an expanded time scale to better show the initial
response to low- (C, left, corresponding
to Bi, right) and high-
(C, right, corresponding to
Bii, right) intensity stimulation.
Recordings in A and B are from different
cells, and traces are single trial examples.
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|
We further characterized the response of stellate cells to parallel
fiber activation with whole-cell current-clamp recordings (Fig.
12B). The effects on stellate cell firing were
similar to those observed in the cell-attached patch configuration. For
low-intensity stimulation, a single stimulus did not trigger an action
potential (Fig. 12Bi, left), but a train
elevated firing rates for several hundred milliseconds (Fig. 12,
Bi, right, C, left). At a
higher intensity, a stimulus train evoked a prolonged depolarization to
approximately 30 mV (Fig. 12, Bii, right,
C, right). The stellate cell did not fire during
this depolarization, likely as a consequence of
Na+ channel inactivation. Similar activity
patterns were observed in all nine cells tested. The duration of this
depolarization is similar to that of the prolonged EPSC recorded in
whole-cell voltage clamp (compare Fig. 12C,
right, with Fig. 1). These findings suggest that that the
prolonged excitatory postsynaptic conductances described in this paper
can interact with voltage-dependent conductances to have profound
effects on stellate cell firing.
| |
DISCUSSION |
Our main finding is that brief stimulus trains can evoke prolonged
synaptic responses mediated by AMPA and NMDA receptors that reflect the
sustained presence of glutamate and glutamate spillover. These results
also highlight the importance of glutamate transporters in regulating
glutamate levels during trains and may have important physiological implications.
EPSCs evoked by single stimuli and trains
The EPSCs evoked by stimulation of a direct pathway with single
stimuli and with brief trains are very different. This reflects both
the contrasting glutamate signals produced by these two stimuli and the
kinetics and glutamate affinities of the activated receptors.
A single stimulus evokes a prominent AMPAR-mediated EPSC (Barbour et
al., 1994; Atluri and Regehr, 1998) and a much smaller NMDAR-mediated
EPSC that is difficult to resolve (Glitsch and Marty, 1999). The brief
EPSC mediated by AMPA receptors is a consequence of a short-lived
glutamate transient that results primarily from phasic release of
neurotransmitter (Atluri and Regehr, 1998; Chen and Regehr, 1999) and
the rapid deactivation kinetics of the AMPA receptors (Barbour et al.,
1994). Furthermore, because the decay time of this rapid AMPAR-mediated
EPSC is unaffected by either an increase in stimulus intensity or the
inhibition of glutamate transporters, it is likely that this glutamate
signal is relatively localized and independent of these transporters
(Barbour et al., 1994). The small, long-lived response mediated by NMDA
receptors is also consistent with a brief glutamate signal activating a small number of NMDA receptors with stereotypically slow kinetics (Lester et al., 1990; Edmonds et al., 1995).
Differences in responses evoked by trains and single stimuli are
apparent for AMPAR- and NMDAR-mediated EPSCs, although these components
are affected in different ways. For synaptic currents mediated by AMPA
receptors, a slow component becomes prominent during trains that is not
apparent in responses to single stimuli (Fig. 3). In contrast, for
EPSCs mediated by NMDA receptors, the most obvious effect of
stimulating with a train is to increase the peak response by 10- to
20-fold, and prolongation of the time course is less evident.
The properties of these EPSCs reflect very different glutamate
signaling during trains. Presynaptic facilitation greatly increases the
release of neurotransmitter during a train (Magleby, 1987; Zucker,
1989; Atluri and Regehr, 1996). Based on the amplitudes of EPSCs
mediated by AMPA receptors, we estimate that, during a train of five
pulses at 100 Hz, facilitation results in a 10- to 20-fold increase in
glutamate release compared with a single stimulus. Trains also
accentuate the delayed release of neurotransmitter, which further
contributes to an increase in transmitter release (Atluri and Regehr,
1998). We propose that the resultant glutamate signal is long-lived and
can spill over from the synaptic cleft.
Several aspects of the responses mediated by AMPA receptors suggest a
sustained elevation of glutamate in responses to trains. First, the
duration of the EPSC persists for longer than can be accounted for by
AMPA receptor kinetics (Barbour et al., 1994; Edmonds et al., 1995).
Second, the prolongation of the EPSC evoked by an increase in stimulus
intensity (Fig. 3) indicates that the glutamate signal becomes larger
and more effective at activating distant AMPA receptors when many
presynaptic fibers are activated. Third, the prolongation of the EPSC
by PDC (Fig. 5) indicates that glutamate transporters normally restrict
the extent of the glutamate signal produced by a stimulus train.
The prominent NMDAR-mediated EPSC evoked by a stimulus train also
reflects both the kinetics of NMDA receptors and the large glutamate
signal generated by the train. The slow kinetics of the NMDA receptors
(Lester et al., 1990; Edmonds et al., 1995) allow the stellate cell to
integrate this glutamate signal over time. Because NMDA receptors have
much slower off rates than AMPA receptors (Edmonds et al., 1995), it is
difficult to resolve differences in the time course of evoked synaptic
responses that are readily apparent with EPSCs mediated by AMPA
receptors. Activation of extrasynaptic NMDA receptors may also
contribute to the properties of the NMDAR-mediated EPSCs (Clark et al.,
1997), although the distribution of NMDA receptors on the stellate cell
is currently unknown.
Glutamate spillover
Our findings establish that glutamate spillover can activate both
NMDA and AMPA receptors at the parallel fiber to stellate cell synapse.
This was first suggested by prolongation of the AMPAR-mediated EPSC by
an increase in stimulus intensity (Fig. 3), as described above. It was
further indicated by the ability to evoke both NMDAR- and
AMPAR-mediated EPSCs with indirect pathway stimulation (Fig. 8).
Moreover, our experiments using MK-801 confirmed that glutamate
spillover can activate NMDA receptors at this synapse (Fig. 9). From
these experiments, we estimate that glutamate released during a
stimulus train can diffuse for tens of micrometers to activate
ionotropic glutamate receptors. It seems likely that this glutamate
spillover contributes to both NMDAR- and AMPAR-mediated EPSCs evoked by
direct pathway stimulation. Finally, our experiments show that these
phenomena occur at both 24 and 34°C.
The AMPAR-mediated EPSCs evoked by direct- and indirect- pathway
stimulus trains indicate that glutamate spillover can activate AMPA
receptors. This result was surprising, because these receptors are
thought to mediate only fast synaptic responses to brief glutamate signals at most synapses. However, previous studies indicate that, at
calyceal and glomerular synapses, glutamate spillover can activate AMPA
receptors (Rossi et al., 1995; Otis and Trussell, 1996; Otis et al.,
1996; Silver et al., 1996; Kinney et al., 1997; Slater et al., 1997;
Overstreet et al., 1999). At these synapses, glutamate is released from
multiple sites into a confined space, which can lead to an extended
glutamate signal and glutamate spillover. However, this specialized
geometry is atypical, and the en passant parallel fiber to
stellate cell synapse is more characteristic of central synapses.
Parallel fibers synapse onto the dendritic shafts of stellate cells at
which bergmann glia processes are sparse (Palay and Chan-Palay, 1974)
and glutamate transporter density is low (Chaudhry et al., 1995). For a
single stimulus, these factors allow for the small amount of released
glutamate to diffuse into the local extrasynaptic space, resulting in a brief glutamate signal (Barbour et al., 1994). However, for a stimulus
train, these same factors permit the large amount of released glutamate
to spill over to nearby synapses. Thus, as at glomerular and calyceal
synapses, the structure of the parallel fiber to stellate cell synapse
also appears to permit AMPA receptor activation by glutamate spillover
in response to a stimulus train.
Significance
Our results may have implications for circuit properties and
plasticities in the cerebellum. Granule cell firing depolarizes both
stellate and Purkinje cells, thereby promoting firing in these two cell
types. As synaptic activation becomes sufficiently large to fire
stellate cells, they will in turn inhibit Purkinje cells and thereby
limit the ability of granule cells to promote Purkinje cell firing.
However, more powerful synaptic activation may prolong the presence of
glutamate at the stellate cell synapse (Fig. 12). The resulting
depolarization could be sufficiently large to prevent stellate cell
firing by inactivating Na+ channels (Fig.
12C, right), thereby reducing inhibition and
increasing the efficacy of parallel fiber inputs to Purkinje cells.
The prolonged EPSCs evoked by high-frequency stimulus trains only
became prominent when large populations of parallel fibers were
activated. This suggests that high-frequency firing of individual granule cells may generate neither an extended glutamate signal nor
spillover. Nevertheless, several points suggest that our results have
broad implications for the behavior of many types of synapses. First,
populations of excitatory neurons, including cerebellar granule cells,
often display high-frequency firing in vivo, particularly during behavioral tasks (Hartmann and Bower, 1998). Second, epilepsy can result in similar high-frequency synchronous firing in neurons. Third, many experiments use similar stimulation protocols to study synaptic transmission and plasticity. Glutamate spillover and prolonged
EPSCs may become prominent in each of these three cases.
| |
FOOTNOTES |
Received Jan. 28, 2000; revised March 17, 2000; accepted March 28, 2000.
This work was supported by National Institutes of Health Grants
R01-NS32405-01 and MH/NS32405-01 to W.G.R. and a National Science
Foundation graduate fellowship to A.G.C. We thank Chinfei Chen, Anatol
Kreitzer, Kaspar Vogt, and Matthew Xu-Friedman for comments on this manuscript.
Correspondence should be addressed to Wade G. Regehr, Department of
Neurobiology, Harvard Medical School, 220 Longwood Avenue, Boston, MA
02115. E-mail: wade_regehr{at}hms.harvard.edu.
| |
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TOP
ABSTRACT
INTRODUCTION
